Download Oxidation of Fatty Acids Is the Source of Increased

Survey
yes no Was this document useful for you?
   Thank you for your participation!

* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project

Document related concepts

Thylakoid wikipedia , lookup

Glycolysis wikipedia , lookup

Fatty acid synthesis wikipedia , lookup

Metabolism wikipedia , lookup

Fatty acid metabolism wikipedia , lookup

Ketosis wikipedia , lookup

Microbial metabolism wikipedia , lookup

Citric acid cycle wikipedia , lookup

Photosynthetic reaction centre wikipedia , lookup

Light-dependent reactions wikipedia , lookup

Evolution of metal ions in biological systems wikipedia , lookup

Reactive oxygen species wikipedia , lookup

Metalloprotein wikipedia , lookup

Free-radical theory of aging wikipedia , lookup

Electron transport chain wikipedia , lookup

Oxidative phosphorylation wikipedia , lookup

Mitochondrial replacement therapy wikipedia , lookup

Mitochondrion wikipedia , lookup

NADH:ubiquinone oxidoreductase (H+-translocating) wikipedia , lookup

Transcript
ORIGINAL ARTICLE
Oxidation of Fatty Acids Is the Source of Increased
Mitochondrial Reactive Oxygen Species Production
in Kidney Cortical Tubules in Early Diabetes
Mariana G. Rosca,1,2 Edwin J. Vazquez,3,4 Qun Chen,5 Janos Kerner,1,2
Timothy S. Kern,2,6 and Charles L. Hoppel1,2,6
Mitochondrial reactive oxygen species (ROS) cause kidney
damage in diabetes. We investigated the source and site of ROS
production by kidney cortical tubule mitochondria in streptozotocininduced type 1 diabetes in rats. In diabetic mitochondria, the increased amounts and activities of selective fatty acid oxidation
enzymes is associated with increased oxidative phosphorylation
and net ROS production with fatty acid substrates (by 40% and
30%, respectively), whereas pyruvate oxidation is decreased and
pyruvate-supported ROS production is unchanged. Oxidation of
substrates that donate electrons at specific sites in the electron
transport chain (ETC) is unchanged. The increased maximal
production of ROS with fatty acid oxidation is not affected by
limiting the electron flow from complex I into complex III. The
maximal capacity of the ubiquinol oxidation site in complex III
in generating ROS does not differ between the control and
diabetic mitochondria. In conclusion, the mitochondrial ETC is
neither the target nor the site of ROS production in kidney
tubule mitochondria in short-term diabetes. Mitochondrial fatty
acid oxidation is the source of the increased net ROS production, and the site of electron leakage is located proximal
to coenzyme Q at the electron transfer flavoprotein that shuttles
electrons from acyl-CoA dehydrogenases to coenzyme Q.
D
iabetic nephropathy (DN) is the leading cause
of end-stage renal disease (ESRD) (1). Hyperlipidemia is an independent factor in renal injury in animals (2) and humans (3). Increased
fatty acid (FA) synthesis enzymes and triglyceride deposition correlated with increased profibrotic factors were
found in the kidney in diabetes in rats (4) and mice (5). Inhibition of lipid synthesis in humans (6) and amelioration
of dyslipidemia (7) protect against diabetic renal disease.
Glomerular disease has been considered the initial and
cardinal manifestation of DN. However, tubulointerstitial
fibrosis is considered a strong predictor of the progressive
loss of renal function leading to ESRD (8,9) and has been
shown to determine the progression to ESRD (10).
From the 1Center of Mitochondrial Diseases, Case Western Reserve University, Cleveland, Ohio; the 2Department of Pharmacology, Case Western Reserve University, Cleveland, Ohio; the 3Department of Nutrition, Case
Western Reserve University, Cleveland, Ohio; the 4Mouse Metabolic Phenotypic Center, Case Western Reserve University, Cleveland, Ohio; the 5Department of Medicine Divisions of Cardiology, Virginia Commonwealth
University, Richmond, Virginia; and the 6Department of Medicine, Case
Western Reserve University, Cleveland, Ohio.
Corresponding author: Mariana G. Rosca, [email protected]
Received 13 October 2011 and accepted 9 March 2012.
DOI: 10.2337/db11-1437
Ó 2012 by the American Diabetes Association. Readers may use this article as
long as the work is properly cited, the use is educational and not for profit,
and the work is not altered. See http://creativecommons.org/licenses/by
-nc-nd/3.0/ for details.
diabetes.diabetesjournals.org
The increase in plasma nonesterified FAs (NEFAs) is a
major component of diabetic dyslipidemia. Tubular cells
are exposed to blood and urine FAs present in free form or
bound to albumin. The expression of cluster of differentiation 36 (CD36) involved in the transport of FA is induced
by high glucose in proximal tubular cells and causes palmitateinduced apoptosis only in human kidneys with diabetic tubular epithelial degeneration (11). Albumin-bound FA, rather
than albumin, itself is a major mediator of tubulointerstitial
lesions in various types of proteinuria-developing ESRD
(12,13).
Proximal tubular cells (.90% of the kidney cortex) engage in active uptake and transepithelial transport of glucose, but only a small amount of glucose, if any, is used
for ATP production (14). Moreover, diabetes causes a
decrease in kidney glucose oxidation due to the inhibition
of pyruvate dehydrogenase activity (15). Skin fibroblasts
isolated from type 1 diabetic patients, with very fast rates
in developing DN, have increased expression of genes involved in mitochondrial FA oxidation (16). The data suggest an energy fuel preference in diabetes favoring fat
oxidation at the expense of glycolysis. The enzyme carnitine
palmitoyltransferase 1 (CPT1) catalyzes the rate-controlling
step in the overall mitochondrial FA b-oxidation pathway.
Its expression is reported to be increased in liver and heart
in diabetes, whereas its sensitivity to malonyl-CoA is reduced,
favoring its activity.
Increased generation of ROS within the mitochondrial
electron transport chain (ETC) has been considered to be
an important contributor to the development of chronic
diabetes complications (17). Increased availability of glucose-generated reducing equivalents in cultured aortic
endothelial (18) and retinal (19) cells or defects in the ETC
in epineural (20) and cardiac (21) mitochondria increase
the electron pressure at specific mitochondrial sites. This
perturbation prolongs the lifetime of partially reduced
intermediates that donate electrons to molecular oxygen
to form superoxide. In contrast, although mitochondrialtargeted antioxidant mitoquinone mesylate (mitoQ) is
beneficial in the treatment of diabetic nephropathy, mitochondrial ETC is not the source of ROS generation in the
kidney of diabetic Akita mice (22) or in the sensory neurons from diabetic rats (23), indicating that there are differences in the source and sites of ROS production in
target organs of diabetes complications. In short-term
diabetes, we found that mitochondrial ETC is neither the
target nor the site of ROS generation in kidney cortical
tubules. The source of increased ROS production is mitochondrial FA b-oxidation; the site of electron leakage is
located proximal to CoQ at the electron transfer flavoprotein that shuttles electrons from acyl-CoA dehydrogenases to coenzyme Q.
Diabetes Publish Ahead of Print, published online May 14, 2012
DIABETES
1
KIDNEY CORTICAL TUBULE MITOCHONDRIA IN DIABETES
RESEARCH DESIGN AND METHODS
Reagents. Unless otherwise specified, all reagents were purchased from
Sigma-Aldrich (St. Louis, MO) and were of the highest purity grade.
Animal model. The investigation conforms to the Guide for the Care and Use
of Laboratory Animals (U.S. National Institutes of Health Publication No. 85-23,
revised 1996). All experimental protocols were approved by the Case Western
Reserve University institutional animal care and use committees. Diabetes was
induced by intraperitoneal injection of streptozotocin (55 mg/kg of body
weight) in citrate buffer to 12-h fasted male Wistar rats. Two units of NPH
insulin were given subcutaneously thrice weekly to avoid severe hypercatabolic state. Glycated hemoglobin was measured by affinity chromatography of the intracardiac blood.
Quantification of serum NEFA and triglyceride-bound glycerol. The
concentration of serum NEFA and of triglyceride-bound glycerol was determined using gas chromatography-electron impact ionization mass spectrometry, as described by Bederman et al. (24). Palmitate, stearate, and oleate
were analyzed as trimethylsilyl derivatives and quantified against a known
amount of heptadecanoic acid. Serum triglyceride-bound glycerol was quantified as a triacetate derivative after an alkaline hydrolysis procedure and the
addition of a known amount of internal standard of deuterium-labeled glycerol.
Isolation of tubule mitochondria. The kidneys cortices were separated from
the medulla. Glomeruli were first isolated (25), and the material consisting of
tubulointerstitium was collected in buffer A (220 mmol/L mannitol, 70 mmol/L
sucrose, 5 mmol/L 3-(N-morpholino)propanesulfonic acid [MOPS], 10 mmol/L
EGTA, 0.2% BSA, pH 7.4). Phase-contrast microscopy was performed during
each experiment to examine the purity of the glomerular material and ensure
that the entire tubular population was harvested within the tubulointerstitial
material. Because the tubules represent the major component of the cortical
tubulointerstitial material, the isolated mitochondria were designated as cortical tubule mitochondria. Tubule mitochondria were isolated (26) and suspended in buffer B (100 mmol/L KCl, 50 mmol/L MOPS, and 0.5 mmol/L EGTA,
pH 7.4).
Oxidative phosphorylation. Oxygen consumption was measured with a
Clark-type electrode (Strathkelvin) in respiration buffer (100 mmol/L KCl, 50
mmol/L MOPS, 1 mmol/L EGTA, 5 mmol/L KH2PO4, and 1 mg/mL defatted BSA,
pH 7.4) (27) at 30°C with glutamate (20 mmol/L) + malate (5 mmol/L), pyruvate (10 mmol/L) + malate, succinate (20 mmol/L) + rotenone (3.75 mmol/L),
durohydroquinone (DHQ, 1 mmol/L) + rotenone, N,N,N9,N9-tetramethyl-pphenylenediamine (TMPD) + ascorbate (1:10 mmol/L) + rotenone, palmitoylCoA (0.02 mmol/L) + carnitine (2 mmol/L) + malate, and palmitoylcarnitine
(0.04 mmol/L) or octanoylcarnitine (0.2 mmol/L) + malate, as substrates. State
3 respiration rate, which is ADP-dependent was measured with 200 mmol/L
ADP. State 4 respiration (ADP-limited) was recorded after ADP consumption.
Then, sequential additions of 2 mmol/L ADP and 200 mmol/L of the uncoupler
dinitrophenol were made to determine the maximal ADP-stimulated and
uncoupled respiratory rates, respectively. State 3-to-state 4 respiratory control
ratios (RCR) reflect the control of oxygen consumption by phosphorylation
(“coupling”). The ADP-to-oxygen ratio (number of ADP molecules added for
each oxygen atom consumed) (28) is an index of the efficiency of oxidative
phosphorylation. In preliminary experiments, the complex IV inhibitor sodium
azide (400 mmol/L) completely inhibited oxidative phosphorylation with
complex IV substrates in both control and diabetic tubule mitochondria,
showing that the azide-sensitive complex IV–supported respiration equals the
TMPD + ascorbate–supported respiration and that there are no differences in
azide sensitivity between control and diabetic tubule mitochondria. Therefore,
the results are presented as TMPD + ascorbate–supported respiration. Rates
of oxygen consumption are expressed in nano atoms (nA) oxygen per minute
per milligram of mitochondrial proteins.
Measurement of CPT1 activity. The activity of CPT1 was determined using
the modified radiochemical forward assay by measuring the formation of 14Clabeled palmitoylcarnitine from 14C-carnitine and palmitoyl-CoA at 37°C as
described (29). CPT1 activity was expressed as the activity inhibited by 200
mmol/L malonyl-CoA and given in nanomoles per minute per milligram of
protein. Malonyl-CoA sensitivity was determined at fixed palmitoyl-CoA concentrations by varying the concentration of malonyl-CoA.
Measurement of fatty acid oxidation enzymes. Frozen-thawed mitochondria were treated with 0.5% cholate and diluted to 0.1 mg/mL buffer A. The
dehydrogenase assays used octanoyl-CoA for medium-chain acyl-CoA dehydrogenase (MCAD) and palmitoyl-CoA for long-chain acyl-CoA dehydrogenase
(LCAD) with phenazine ethosulfate as an intermediate electron acceptor from
flavin adenine dinucleotide (FADH2) and oxidized cytochrome c as the final
electron acceptor. The reduction of cytochrome c was recorded spectrophotometrically at 550 nm.
Western blot analysis. Proteins (equal amounts of kidney tubule mitochondrial protein [18 mg/lane], heart mitochondrial protein [6.5 mg/lane], and
liver outer membrane protein [0.5 mg/lane]) were subjected to SDS-PAGE,
2
DIABETES
electroblotted onto polyvinylidene fluoride membranes, and probed with
specific antibodies. Densitometric evaluation of the gels was carried out using
ImageJ software (National Institutes of Health) coupled with Multi Gauge V3.0
(Fujifilm Life Science).
The antibodies used in this study were as follows: liver ( 620MDPKSTAEQRLK631)
and liver + muscle (230NYVSDWWEEYIYLRGR245) isoforms of CPT1 (30) at
1:3000 dilutions in overnight incubations; beef liver mitochondrial CPT2 (29),
carnitine acyl-carnitine translocase (CACT) (31), MCAD (Epitomics) at 1:3000
dilutions, each in 2-h successive incubations; manganese superoxide dismutase (Mn-SOD) and copper/zinc SOD (Cu/Zn-SOD; Assay Designs) at 1:5000
dilutions in overnight incubations. All membranes were stripped and incubated with iron sulfur protein antibody (MitoSciences) as a loading control
at 1:5000 dilution overnight. The amount of this protein is unchanged in kidney
mitochondria in type 1 diabetes (32). Secondary antibodies (BioRad) were
used at 1:10000 dilutions in 1 h incubation. Multiple exposure times were used
for developing, and the optimal ones were chosen for presentation.
Measurement of hydrogen peroxide (H2O2) produced by mitochondria.
The production of ROS by intact mitochondria was measured as the rate of
oxidation of the fluorogenic indicator, Amplex red, by H2O2 in the presence of
horseradish peroxidase (33). H2O2 production was initiated by substrates in
concentrations similar to those indicated for oxidative phosphorylation measurements. Rotenone and antimycin A were added to inhibit the activities of
complex I and III, respectively. Fluorescence was recorded in a microplate
reader with 530-nm excitation and 590-nm emission wavelengths. Standard
curves with known amounts of H2O2 were linear up to 2 mmol/L H2O2. The rate
of H2O2 production also was linear with respect to mitochondrial protein. The
results are presented as fluorescence in the presence of mitochondria minus
the background (absence of mitochondria) in picomoles of H2O2 per milligram
of mitochondrial protein/30 min.
Statistical analysis. The results were analyzed using the Student two-tailed
two-sample t test. Data are reported as mean 6 SEM. P , 0.05 was considered
significant.
RESULTS
Clinical and biochemical characteristics of diabetic
rats. Diabetic rats did not lose weight during the study but
showed decreased weight gain, hyperglycemia, polyuria,
and increased glycated hemoglobin levels 8–9 weeks after
the onset of diabetes (Table 1). The serum was macroscopically turbid, indicating secondary diabetic hyperlipidemia. Glycerol and NEFA levels were increased in sera
of diabetic rats compared with the control rats. Kidney
weight was significantly increased in the diabetic group,
suggesting renal hypertrophy. Mitochondrial yield does not
change in diabetes (20.1 6 2.9 vs. 18.9 6 1.2 mg mitochondrial protein/g wet cortex tissue in control and diabetic
TABLE 1
Clinical and biochemical characteristics of control and diabetic
rats
Study duration (weeks)
Body weight (g)
Beginning of the study
End of the study
Glycemia (mg/100 mL)
Glycated Hb (HbA1c, %)
Serum visual appearance
Glycerol (mmol/L)
Total NEFA (mmol/L)
Palmitic (mmol/L)
Oleic (mmol/L)
Stearic (mmol/L)
Kidney weight (g)
Diuresis (mL/day)
Control rats
(n = 7)
Diabetic rats
(n = 6)
9.0 6 1.6
8.0 6 1.0
310 6 12
410 6 24
142.5 6 10.5
2.9 6 0.1
Clear
2.114 6 0.169
0.370 6 0.007
0.156 6 0.003
0.095 6 0.004
0.119 6 0.008
1.5 6 0.0
11.8 6 1.5
275 6 15
320 6 37*
255.1 6 4.5*
9.7 6 0.3*
Turbid
19.208 6 2.109*
1.256 6 0.131*
0.490 6 0.046*
0.421 6 0.046*
0.345 6 0.034*
1.8 6 0.1*
153.0 6 3.6*
Data are presented as mean 6 SEM. *P , 0.05.
diabetes.diabetesjournals.org
M.G. ROSCA AND ASSOCIATES
tubules, respectively), suggesting that there has been no
change in mitochondrial content in kidney tubules.
Substrate processing rather than the mitochondrial
ETC is changed in kidney tubules in diabetes. Diabetic
tubule kidney mitochondria oxidizing glutamate have
higher state 3 respiratory rates compared with the control
(Table 2). The ADP-to-oxygen ratios were unchanged,
showing that the coupling efficiency of the tubule kidney
mitochondria is preserved in diabetes. With the uncoupler,
dinitrophenol, the glutamate-supported respiratory rates of
nondiabetic tubule mitochondria do not increase nor do
they reach the level of respiratory rates of diabetic mitochondria (Table 2). The oxidation of glutamate + malate is
not changed by diabetes in tubule mitochondria, whereas
pyruvate + malate oxidation is lower in diabetic tubule
mitochondria compared with the control (Fig. 1A). The
oxidation of succinate, which donates electrons to FAD in
complex II, was unchanged in diabetic tubule mitochondria, as were the state 3 respiratory rates with durohydroquinone (complex III electron donor) and those with the
cytochrome c electron donor TMPD + ascorbate (Fig. 1A).
The data show that the ETC complexes in tubule mitochondria are not the targets of diabetes-induced alterations
in this experimental model.
Mitochondrial FA oxidation is increased in kidney
tubules in diabetes. CPT1 is the rate-controlling enzyme
in mitochondrial FA oxidation. We measured oxidative
phosphorylation rates with CPT1-dependent and CPT1independent substrates. Palmitoyl-CoA is converted to palmitoylcarnitine by the outer membrane CPT1. Palmitoylcarnitine
and octanoylcarnitine are transported into the mitochondrial matrix via the CACT bypassing CPT1, and react with
CoA in the matrix in a reaction catalyzed by the inner
membrane CPT2. Within the matrix, palmitoyl-CoA and
octanoyl-CoA undergo sequential b oxidation, producing
acetyl-CoA, NADH, and FADH2. Electrons from NADH
enter the ETC at complex I, whereas those from the FADH2
of acyl-CoA dehydrogenases enter the ETC at complex III
via electron transfer flavoprotein (ETF) and ETF-coenzyme
Q oxidoreductase (ETF-QOR). As shown in Fig. 1B, CPT1dependent and CPT1-independent lipid substrates were
both oxidized at higher rates by diabetic tubule mitochondria.
CPT1 activity is not changed by diabetes despite the
increased protein amount. CPT1 converts long-chain
acyl-CoA to acylcarnitines and exists as two isoforms, the
muscle (CPT1b) and the liver (CPT1a) isoforms, which
TABLE 2
Oxidative properties of kidney cortical tubule mitochondria
isolated from kidneys of control and diabetic rats with
glutamate as the substrate
Control rats
n=7
State 3
State 4
RCR
ADP-to-oxygen ratio
Maximal ADP
Dinitrophenol
100.0
22.0
5.2
3.1
110.0
80.0
6
6
6
6
6
6
8.3
3.7
0.7
0.2
9.9
3.8
Diabetic rats
n=6
154.0
20.0
8.1
3.0
159.0
128.0
6
6
6
6
6
6
12.9*
1.6
0.8*
0.2
12.8*
14.8*
Data are presented as mean 6 SEM. State 3 was induced with 0.2
mmol/L ADP. Respiratory rates are expressed as nA O/min/mg mitochondrial protein. Maximal ADP: State 3 with saturating 2 mmol/L
ADP. *P , 0.05.
diabetes.diabetesjournals.org
differ with respect to their kinetic properties. The activity
of CPT1 was not changed by diabetes in kidney tubule
mitochondria (Fig. 2A). The in vivo flux through CPT1
depends on the sensitivity of the enzyme to malonyl-CoA
inhibition. Liver mitochondria are significantly less sensitive to malonyl-CoA inhibition (half-maximal inhibitory
concentration [IC50] ;5 mmol/L) compared with kidney
tubular mitochondria (IC50 ;2 mmol/L; Fig. 2B). Figure 2C
shows that a slight but significantly higher percentage of
CPT I is not inhibited by 2 mmol/L malonyl-CoA (equals the
IC50 for nondiabetic mitochondria) in diabetic tubule mitochondria compared with the control, suggesting a decrease in the sensitivity of CPT1 to malonyl-CoA in kidney
tubules in diabetes.
The two CPT1 isoforms show differences in migration
(Fig. 2D), with slightly higher calculated size of CPT1a
(;88 kDa) than CPT1b (;82 kDa), as reported (34).
CPT1b protein predominates in the heart mitochondria,
whereas only CPT1a is present in liver mitochondrial outer
membrane. CPT1a is the only isoform expressed in kidney
tubule mitochondria, and the amount of CPT1a protein is
increased in diabetes.
The activities and amounts of selective FA b-oxidation
enzymes are increased in kidney tubules in diabetes.
Figure 3A shows the increase in the activity of mediumchain and long-chain acyl-CoA dehydrogenases in tubule
mitochondria isolated from the diabetic group. The specific activity of citrate synthase, a mitochondrial marker
enzyme, does not differ in tubule mitochondria isolated
from the control or diabetic group, indicating that the
purity of both preparations was identical. The amounts of
CACT and CPT2 proteins are increased in tubule mitochondria isolated from diabetic kidneys compared with the
controls (Fig. 3B).
Increased capacity of diabetic tubule mitochondria to
produce ROS is supported by oxidation of FA substrates;
the potential site of electron leakage is located proximal
to coenzyme Q within the FA oxidation pathway. When
added to intact mitochondria, Amplex red detects the net
mitochondrial release of H2O2 into the buffer when H2O2
production by SOD surpasses its consumption by mitochondrial antioxidant defense mechanisms. Superoxide generated toward and within the mitochondrial matrix is
converted to H2O2 by the matrix Mn-SOD, whereas the superoxide generated toward the intermembrane space is
converted to H2O2 by the intermembrane Cu/Zn-SOD.
Surprisingly, control and diabetic tubule mitochondria
both produce equal amounts of H2O2 when oxidizing pyruvate (Fig. 4A), despite the decreased oxidation of pyruvate in diabetes (Fig. 1). Rotenone (complex I inhibitor that
limits the electron flow into complex III) does not additionally increase but rather decreases the release of H2O2.
The addition of antimycin A has no effect on H2O2 in the
control or diabetic tubule mitochondria oxidizing pyruvate.
In contrast, diabetic tubule mitochondria oxidizing a FA
substrate (Fig. 4B) produce a larger amount of H2O2 than
the control mitochondria under basal conditions (no inhibitor added). Rotenone does not additionally increase
but rather decreases the H2O2 production in control and
diabetic tubule mitochondria. In contrast, the complex III
inhibitor antimycin A significantly increases the production
of H2O2 in diabetic tubule mitochondria but has no effect
on control tubule mitochondria. The data indicate that the
maximal capacity (antimycin A-added) of kidney tubule
mitochondria oxidizing fat substrates is increased by diabetes; rotenone does not decrease this H2O2 maximal
DIABETES
3
KIDNEY CORTICAL TUBULE MITOCHONDRIA IN DIABETES
FIG. 1. State 3 respiratory rates of kidney cortical tubule mitochondria. A: ETC site-specific substrates. Substrates that donate the reducing
equivalents to complex I (G+M, glutamate + malate; P+M, pyruvate + malate), II (S, succinate + rotenone), III (DHQ, DHQ + rotenone), and IV
(TMPD+Asc, TMPD + ascorbate + rotenone) were used. B: Lipid substrates. PC+M, palmitoylcarnitine + malate; P+C+M, palmitoyl-CoA + carnitine +
malate; OC+M, octanoylcanitine + malate. State 3 was induced with 0.2 mmol/L ADP. *P < 0.05 control (n = 7) vs. diabetic (n = 6). Mean 6 SEM.
release capacity with fat substrates. H2O2 released by
control and diabetic tubule mitochondria oxidizing pyruvate or fat substrate was dissipated by catalase at similar
extent, indicating that the increase in the measured fluorescence was because of H2O2 and not induced by the
chemical reaction of Amplex red with FA hydroperoxides.
When oxidizing succinate at complex II (Fig. 4C), control and diabetic tubule mitochondria both produce H2O2,
which was decreased in both by ;45% with rotenone,
which inhibits the reversed electron transport toward
complex I. The data indicate that a major source of superoxide in kidney tubule mitochondria oxidizing succinate is the electron back-flow to complex I and that this
feature is unchanged by diabetes. The addition of antimycin A increases the production of H2O2 in control and
diabetic tubule mitochondria oxidizing succinate.
This increase in the net release of H2O2 by diabetic mitochondria with fat substrates indicates that the production
surpasses its consumption by mitochondrial antioxidant
defense mechanisms. The amount of both SODs determined
by immunoblot analysis was not significantly changed by
diabetes (Fig. 4D). These data indicate that it is the production of superoxide rather than its dismutation that leads
to increased H2O2 release in the buffer by diabetic kidney
tubule mitochondria.
DISCUSSION
This study shows that 8- to 9-week type 1 diabetes induces
an increase in mitochondrial FA b-oxidation without defects
4
DIABETES
in the electron transport and identifies oxidation of FA
rather than glycolysis-derived substrate (pyruvate) oxidation as the source of mitochondrial ROS in diabetic
tubules. Because more than 90% of kidney cortex consists of
proximal tubules, our results can be attributed to proximal
tubule mitochondria. These changes in mitochondria coexist with renal structural and functional modifications
consistent with the early stage of DN (35,36).
Diabetic cortical tubule mitochondria reach maximal
respiratory rates with glutamate when oxidation is coupled
with and depends on phosphorylation of ADP, and do not
further increase when the control of oxidation by phosphorylation is eliminated by the uncoupler. These data
show that the control site of oxidative phosphorylation is
located at the level of glutamate oxidation to form NADH
or electron transport rather than at the level of the phosphorylation system. When substrates that donate electrons
at specific sites in the ETC were used, no change in respiratory rates was detected, showing that the formation of
NADH from glutamate (glutamate transporter or dehydrogenase) rather than the electron transport is increased in
diabetes.
The decrease in pyruvate + malate oxidation in diabetic
tubule mitochondria suggests a decrease in either pyruvate
transporter or pyruvate dehydrogenase that was reported
inhibited in the diabetic kidney (15,37). However, the oxidation of pyruvate in isolated tubule mitochondria was
performed with a concentration of pyruvate (10 mmol/L)
that keeps pyruvate dehydrogenase in its active form
diabetes.diabetesjournals.org
M.G. ROSCA AND ASSOCIATES
FIG. 2. CPT1 in kidney cortical tubule mitochondria. A: Specific activity of CPT1 in tubule mitochondria from control and diabetic kidneys.
B: Comparison of malonyl-CoA sensitivity (IC50) of CPT1 in mitochondria from control rat liver and kidney cortical tubules. C: The percentage of
CTP1 not inhibited by 2 mmol/L malonyl-CoA in tubule mitochondria from control and diabetic kidneys. D: Semiquantitative determination CPT1
protein with antibodies prepared against the amino acid sequence is shown. The bar graphs represent the densitometric quantitation of the
immunoblots expressed in arbitrary units normalized per iron sulfur protein (ISP). H, heart subsarcolemmal mitochondria; L, liver mitochondrial
outer membrane. *P < 0.05 control vs. diabetic. Mean 6 SEM. (A high-quality color representation of this figure is available in the online issue.)
diabetes.diabetesjournals.org
DIABETES
5
KIDNEY CORTICAL TUBULE MITOCHONDRIA IN DIABETES
FIG. 3. The specific activities and immunoquantitation of fatty acid b-oxidation enzymes in kidney cortical tubule mitochondria. A: Enzyme activities were measured in detergent-solubilized, frozen-thawed mitochondria from control and diabetic rats. CS, citrate synthase. B: Immunoblots
and densitometric quantitation of the immunoblots are expressed in arbitrary units normalized per iron sulfur protein (ISP). *P < 0.05 control vs.
diabetic. Mean 6 SEM.
by inhibiting pyruvate kinase (38). Therefore, pyruvate
transporter rather than pyruvate dehydrogenase seems to
be responsible for the decrease in pyruvate oxidation in
diabetic tubule mitochondria.
6
DIABETES
Diabetic tubule mitochondria oxidize FA substrates at
higher rates than the controls. This is associated with an
increase in the specific activities of both MCAD and
LCAD without changes in the MCAD amount, suggesting
diabetes.diabetesjournals.org
M.G. ROSCA AND ASSOCIATES
FIG. 4. Kidney cortical tubule mitochondria production of H2O2 with pyruvate + malate (A) palmitoylcarnitine + malate (B), and succinate +
rotenone (C) as the substrates. Rot, rotenone; AA, antimycin A; Cat, catalase. D: Semiquantitative determination of mitochondrial Mn-SOD and
Cu/Zn-SOD. The bar graphs represent the densitometric quantitation of the immunoblots expressed in arbitrary units normalized per iron sulfur
protein (ISP). H, heart subsarcolemmal mitochondria. *P < 0.05 compared with the indicated groups. Mean 6 SEM. (A high-quality color representation of this figure is available in the online issue.)
diabetes.diabetesjournals.org
DIABETES
7
KIDNEY CORTICAL TUBULE MITOCHONDRIA IN DIABETES
post-translational modifications that stimulate its catalytic
activity. In contrast, despite the increase in CPT1a protein
amount that favors mitochondrial FA b-oxidation, the catalytic activity of CTP1a is unchanged in diabetes, suggesting
that CPT1 has a low flux control coefficient for mitochondrial FA b-oxidation in kidney tubules in diabetes.
The twofold higher IC50 for malonyl-CoA of CPT1
assessed in control tubule mitochondria compared with
liver mitochondria is not due to the presence of the CPT1b
isoform, as shown by the Western blot approach. We
propose that this kinetic property of CPT1a is an intrinsic
feature of this isoform in kidney tubule mitochondria due
to differences in outer membrane lipid environment or to
post-translational modifications.
Proximal kidney tubules (.90% of the kidney cortex)
rely on FA rather than glucose to form ATP during normal
metabolic conditions (14); we found that this feature is
enhanced in diabetes. Peroxisome proliferator-activated
receptors (PPAR) are potential candidates that induce the
enhanced mitochondrial FA b-oxidation in kidney tubules
in diabetes. All members of this subfamily of nuclear
receptors were detected in kidney cortex (39), are activated by NEFA (40), which were found increased in the
diabetic sera in our model (Table 1), induce expression of
FA b-oxidation enzymes (41), and promote FA utilization.
Our data also show that diabetic tubule mitochondria
produce more ROS than the controls under both basal and
maximal conditions when oxidizing FA substrates compared with oxidation of pyruvate. Prior reports suggesting
that pyruvate may be the source of ROS (42) were based
on the association of the elevated inner membrane potential and ROS with the increased content of both pyruvate and the tricarboxylic cycle intermediates originating
from the pyruvate carboxylase route in mitochondria isolated from the whole kidney tissue. Our results contradict
this assumption. Owing to kidney cellular complexity, the
current study was performed specifically on mitochondria
isolated from cortical tubular structures (highly oxidative
nephron segments that depend on the ATP generated from
FA oxidation) free of medullary structures (highly glycolytic). The inhibition of the pyruvate dehydrogenase in the
diabetic medulla may lead to the increased content of pyruvate within mitochondria isolated from the whole kidney.
Oxidation of FA substrates was reported to be a source
of ROS in the normal heart (43) and skeletal muscle (44)
mitochondria. Increased oxidation of FA in aortic endothelial cells was reported to increase superoxide production from sites located within the electron transport
(45,46). Fatty acids are complex I and III substrates via
NADH and FADH2–ETF/ETF-QOR, respectively (Fig. 5).
Rotenone blocks the distal part of complex I, increases the
reduction of the NADH dehydrogenase site of complex I,
and causes an electron leak toward the matrix in cardiac
mitochondria (33,43). Because rotenone does not increase
H2O2 production with lipid and nonlipid complex I substrates in control and diabetic tubule mitochondria, we
propose that complex I is not the main site of ROS production in tubule mitochondria or that complex I–generated
oxidants are directed toward the mitochondrial matrix and
inactivated by matrix-antioxidant systems in both control
and diabetic states. The equal decrease of ROS with rotenone indicates that an additional site rather than the reverse electron transport to complex I is responsible for the
increased ROS generation by diabetic tubule mitochondria
oxidizing fat substrates.
8
DIABETES
Complex III is a key site for ROS generation in cardiac
mitochondria (33,43). Because the limitation of the electron flow from complex I into complex III with rotenone
equally decreased the release of H2O2 by control and diabetic mitochondria oxidizing pyruvate, the data show that
complex III is the sole contributor to the release of H2O2
by tubular mitochondria oxidizing pyruvate. In contrast
with cultured cells (45,46), the rotenone-inhibited tubule
mitochondria from diabetic kidney show a higher amount
of H2O2 than the control mitochondria oxidizing fat substrates. These data also indicate that a site other than
complex III significantly contributes to ROS generation
when fatty acids are used as substrates in diabetes.
The ubiquinol oxidation (Qo) site in complex III is the
main site of electron leak oriented equally toward the intermembrane space and mitochondrial matrix (47). Antimycin A binds to complex III at a site that overlaps the
ubiquinone reduction (Qi) site, increases electron leak
from the Qo site, and increases superoxide generation. The
minimal increase in H2O2 production in control mitochondria with antimycin A indicates that the superoxide
production is oriented mainly toward the matrix and
scavenged by the matrix antioxidants. The different topology of the superoxide released by kidney tubule mitochondria compared with cardiac mitochondria may reside
in differences in lipid structure of the inner membrane
responsible for a different mobility of the semiquinone in
the Q cycle.
Antimycin A increases H2O2 released by diabetic tubule
mitochondria oxidizing only fat substrates, indicating that
major changes in the topology of ROS production are induced by diabetes in kidney tubule mitochondria when
oxidizing fat. With succinate + rotenone + antimycin, diabetic tubule mitochondria do not show a significantly
higher maximal capacity to produce H2O2 compared with
the control mitochondria (P = 0.081). Because complex II
is not considered to be the major site of superoxide production, the data suggest that the maximal capacity of the
Qo site in complex III to generate oxidants does not differ
between the control and diabetic tubule mitochondria.
A more likely explanation is that an additional site
within the ETF/ETF-QOR–Qo segment (Fig. 5) becomes a
site of superoxide production. The first dehydrogenase
reaction in FA b-oxidation generates FADH2, with the
electron donated to ETF. The FAD prosthetic group of
ETF is rapidly reduced to the semiquinone form and more
slowly to the fully reduced form (48,49), suggesting that
the semiquinone form of the ETF may be the electron
donor for the univalent reduction of oxygen to form superoxide. Similarly, the FAD prosthetic group of ETF-QOR
cycles between the oxidized and partially reduced (semiquinone) forms. However, crystallography studies of ETFQOR show that the semiquinone is protected from reacting
with molecular oxygen (50) and that ETF-QOR is unlikely
to be a site of superoxide production. Specific inhibitors
must be developed to evaluate the real contribution of ETF
or ETF-QOR to oxyradical production.
The matrix-generated superoxide from the site located
at the ETF is dismutated to H2O2 by the matrix Mn-SOD.
This enzyme reroutes superoxide from combining with
nitric oxide to form peroxynitrite, which may alter mitochondrial function via tyrosine nitration of the ETC protein
subunits. Therefore, the fate of mitochondrial ETC in longterm diabetes is controlled by crosstalk between mitochondrial antioxidant defense mechanisms.
diabetes.diabetesjournals.org
M.G. ROSCA AND ASSOCIATES
FIG. 5. The passage of electrons from FA to the ETC. Sites of electron leak and superoxide production and its dismutation to H2O2 are shown. The
electron passage is shown in red. Electrons are released by substrate oxidation and captured by NADH and FADH2, which are oxidized by complexes I and II, respectively. These complexes pass the electrons to ubiquinone (Q). The reduced Q (ubiquinol, QH2) diffuses to and binds within
the QH2 oxidation site (Qo) in complex III (formed by cytochrome b and the Fe-S protein rotated toward it). ETF linked to ETF-QOR provides an
alternative pathway to pass electrons to the ETC. The FAD cofactors in ETF and ETF-QOR carry 1 electron at a time with the formation of intermediate semiquinone state of flavin. Flavin in the ETF is rapidly reduced by one electron by acyl-CoA dehydrogenases (ACAD). The slow
complete reduction of flavin facilitates the accumulation of semiquinone. The electrons are donated sequentially to the Fe-S center and flavin in
the ETF-QOR. Both electrons are further donated to Q (bound to the ETF-QOR in the mitochondrial inner membrane). The QH2 product leaves this
site and is reoxidized in the Qo center of complex III. The transfer of electrons diverges within the Qo site: one electron is transferred to the Fe-S,
followed immediately by the other electron transfer to the Qi (Q reduction site) via cytochrome bL. Under conditions of uninhibited electron flow
through complex III, the reaction is rapid, preventing the accumulation of semiquinone (QH∙) in the Qo site. Antimycin A, a Qi site inhibitor,
prevents the electron from leaving complex III, leading to accumulation of QH∙ and O2∙ formation. Rotenone, an inhibitor of the Q reduction site in
complex I, prevents the electrons from leaving complex I and reaching complex III, leading to an increase in O2∙ formation from complex I toward
the matrix and decrease in O2∙ formation from Qo in complex III toward the intermembrane space. Superoxide is dismutated by the matrix Mn-SOD
and intermembrane space Cu/Zn-SOD. (A high-quality color representation of this figure is available in the online issue.)
In conclusion, mitochondrial ETC is neither the target nor
the ROS-generating site in short-term diabetes. This study
identifies FA as the source of reducing equivalents responsible for increased ROS production by kidney tubule
mitochondria in diabetes and shows that ETF is the major
site of electron leakage. A future time course study will
help to link the increase in ROS production originating
from mitochondrial FA oxidation pathway with the ETC
damage and the progression to advanced stages of diabetic
kidney disease.
ACKNOWLEDGMENTS
Funding for this work was provided by the National
Institutes of Health Grant EU-00300 (to T.S.K.) and DK066107 (to C.L.H.).
No potential conflicts of interest relevant to this article
were reported.
M.G.R. formulated the hypothesis, researched and interpreted the data, and wrote the manuscript. E.J.V. and Q.C.
researched data. J.K. researched data, contributed to discussion and interpretation of the data, and reviewed the
manuscript. T.S.K. and C.L.H. contributed to discussion and
interpretation of the data and reviewed the manuscript.
M.G.R. and C.L.H. are the guarantors of this work and, as
such, had full access to all the data in the study and take
responsibility for the integrity of the data and the accuracy
of the data analysis.
The authors acknowledge the Mouse Metabolic Phenotypic
Center at Case Western Reserve University for performing
diabetes.diabetesjournals.org
the measurements of glycerol and NEFA levels. The
authors thank Dr. Bernard Tandler (School of Dental
Medicine, Case Western Reserve University) and the
Hoppel Laboratory “Writing with Style” group for editorial
assistance.
REFERENCES
1. USRDS. USRDS: the United States Renal Data System. Am J Kidney Dis
2003;42(6 Suppl. 5):1–230
2. Spencer MW, Mühlfeld AS, Segerer S, et al. Hyperglycemia and hyperlipidemia act synergistically to induce renal disease in LDL receptordeficient BALB mice. Am J Nephrol 2004;24:20–31
3. Kambham N, Markowitz GS, Valeri AM, Lin J, D’Agati VD. Obesityrelated glomerulopathy: an emerging epidemic. Kidney Int 2001;59:1498–
1509
4. Sun L, Halaihel N, Zhang W, Rogers T, Levi M. Role of sterol regulatory
element-binding protein 1 in regulation of renal lipid metabolism and
glomerulosclerosis in diabetes mellitus. J Biol Chem 2002;277:18919–
18927
5. Proctor G, Jiang T, Iwahashi M, Wang Z, Li J, Levi M. Regulation of renal
fatty acid and cholesterol metabolism, inflammation, and fibrosis in Akita
and OVE26 mice with type 1 diabetes. Diabetes 2006;55:2502–2509
6. Fried LF, Orchard TJ, Kasiske BL. Effect of lipid reduction on the progression of renal disease: a meta-analysis. Kidney Int 2001;59:260–269
7. Nakamura T, Kawagoe Y, Ogawa H, et al. Effect of low-density lipoprotein
apheresis on urinary protein and podocyte excretion in patients with nephrotic syndrome due to diabetic nephropathy. Am J Kidney Dis 2005;45:
48–53
8. Lane PH, Steffes MW, Fioretto P, Mauer SM. Renal interstitial expansion in
insulin-dependent diabetes mellitus. Kidney Int 1993;43:661–667
9. White KE, Bilous RW. Type 2 diabetic patients with nephropathy show
structural-functional relationships that are similar to type 1 disease. J Am
Soc Nephrol 2000;11:1667–1673
DIABETES
9
KIDNEY CORTICAL TUBULE MITOCHONDRIA IN DIABETES
10. Bohle A, Wehrmann M, Bogenschütz O, Batz C, Müller CA, Müller GA. The
pathogenesis of chronic renal failure in diabetic nephropathy. Investigation of 488 cases of diabetic glomerulosclerosis. Pathol Res Pract
1991;187:251–259
11. Susztak K, Ciccone E, McCue P, Sharma K, Böttinger EP. Multiple metabolic hits converge on CD36 as novel mediator of tubular epithelial apoptosis in diabetic nephropathy. PLoS Med 2005;2:e45
12. Thomas ME, Morrison AR, Schreiner GF. Metabolic effects of fatty acidbearing albumin on a proximal tubule cell line. Am J Physiol 1995;268:
F1177–F1184
13. Ghiggeri GM, Ginevri F, Candiano G, et al. Characterization of cationic
albumin in minimal change nephropathy. Kidney Int 1987;32:547–553
14. Guder WG, Wagner S, Wirthensohn G. Metabolic fuels along the nephron:
pathways and intracellular mechanisms of interaction. Kidney Int 1986;29:
41–45
15. Guder WG, Schmolke M, Wirthensohn G. Carbohydrate and lipid metabolism of the renal tubule in diabetes mellitus. Eur J Clin Chem Clin Biochem 1992;30:669–674
16. Huang C, Kim Y, Caramori ML, et al. Diabetic nephropathy is associated
with gene expression levels of oxidative phosphorylation and related
pathways. Diabetes 2006;55:1826–1831
17. Brownlee M. The pathobiology of diabetic complications: a unifying
mechanism. Diabetes 2005;54:1615–1625
18. Nishikawa T, Edelstein D, Du XL, et al. Normalizing mitochondrial superoxide production blocks three pathways of hyperglycaemic damage.
Nature 2000;404:787–790
19. Du Y, Miller CM, Kern TS. Hyperglycemia increases mitochondrial superoxide in retina and retinal cells. Free Radic Biol Med 2003;35:1491–1499
20. CoppeyLJ, Gellett JS, Davidson EP, Yorek MA. Preventing superoxide formation in epineurial arterioles of the sciatic nerve from diabetic rats restores endothelium-dependent vasodilation. Free Radic Res 2003;37:33–40
21. Lashin OM, Szweda PA, Szweda LI, Romani AM. Decreased complex II
respiration and HNE-modified SDH subunit in diabetic heart. Free Radic
Biol Med 2006;40:886–896
22. Chacko BK, Reily C, Srivastava A, et al. Prevention of diabetic nephropathy in Ins2(+/)⁻(AkitaJ) mice by the mitochondria-targeted therapy MitoQ.
Biochem J 2010;432:9–19
23. Akude E, Zherebitskaya E, Chowdhury SK, Smith DR, Dobrowsky RT,
Fernyhough P. Diminished superoxide generation is associated with respiratory chain dysfunction and changes in the mitochondrial proteome of
sensory neurons from diabetic rats. Diabetes 2011;60:288–297
24. Bederman IR, Foy S, Chandramouli V, Alexander JC, Previs SF. Triglyceride synthesis in epididymal adipose tissue: contribution of glucose
and non-glucose carbon sources. J Biol Chem 2009;284:6101–6108
25. Kreisberg JI, Hoover RL, Karnovsky MJ. Isolation and characterization of
rat glomerular epithelial cells in vitro. Kidney Int 1978;14:21–30
26. Rosca MG, Mustata TG, Kinter MT, et al. Glycation of mitochondrial proteins from diabetic rat kidney is associated with excess superoxide formation. Am J Physiol Renal Physiol 2005;289:F420–F430
27. Hoppel C, DiMarco JP, Tandler B. Riboflavin and rat hepatic cell structure
and function. Mitochondrial oxidative metabolism in deficiency states.
J Biol Chem 1979;254:4164–4170
28. Chance B, Williams GR. Respiratory enzymes in oxidative phosphorylation. III. The steady state. J Biol Chem 1955;217:409–427
29. Hoppel CL, Kerner J, Turkaly P, Turkaly J, Tandler B. The malonyl-CoAsensitive form of carnitine palmitoyltransferase is not localized exclusively
in the outer membrane of rat liver mitochondria. J Biol Chem 1998;273:
23495–23503
30. Distler AM, Kerner J, Lee K, Hoppel CL. Post-translational modifications
of mitochondrial outer membrane proteins. Methods Enzymol 2009;457:
97–115
10
DIABETES
31. Hoppel C, Kerner J, Turkaly P, Tandler B. Rat liver mitochondrial contact
sites and carnitine palmitoyltransferase-I. Arch Biochem Biophys 2001;392:
321–325
32. Bugger H, Chen D, Riehle C, et al. Tissue-specific remodeling of the
mitochondrial proteome in type 1 diabetic akita mice. Diabetes 2009;58:
1986–1997
33. Chen Q, Vazquez EJ, Moghaddas S, Hoppel CL, Lesnefsky EJ. Production
of reactive oxygen species by mitochondria: central role of complex III.
J Biol Chem 2003;278:36027–36031
34. McGarry JD, Brown NF. The mitochondrial carnitine palmitoyltransferase
system. From concept to molecular analysis. Eur J Biochem 1997;244:1–14
35. Liu CX, Hu Q, Wang Y, et al. Angiotensin-converting enzyme (ACE) 2
overexpression ameliorates glomerular injury in a rat model of diabetic
nephropathy: a comparison with ACE inhibition. Mol Med 2011;17:59–69
36. Sato Y, Feng GG, Huang L, et al. Enhanced expression of naofen in kidney
of streptozotocin-induced diabetic rats: possible correlation to apoptosis
of tubular epithelial cells. Clin Exp Nephrol 2010;14:205–212
37. Huang B, Wu P, Popov KM, Harris RA. Starvation and diabetes reduce the
amount of pyruvate dehydrogenase phosphatase in rat heart and kidney.
Diabetes 2003;52:1371–1376
38. Chiang PK, Sacktor B. Control of pyruvate dehydrogenase activity in intact
cardiac mitochondria. Regulation of the inactivation and activation of the
dehydrogenase. J Biol Chem 1975;250:3399–3408
39. Wang YX. PPARs: diverse regulators in energy metabolism and metabolic
diseases. Cell Res 2010;20:124–137
40. Bensinger SJ, Tontonoz P. Integration of metabolism and inflammation by
lipid-activated nuclear receptors. Nature 2008;454:470–477
41. Ouali F, Djouadi F, Merlet-Bénichou C, Bastin J. Dietary lipids regulate
beta-oxidation enzyme gene expression in the developing rat kidney. Am
J Physiol 1998;275:F777–F784
42. de Cavanagh EM, Ferder L, Toblli JE, et al. Renal mitochondrial impairment is attenuated by AT1 blockade in experimental Type I diabetes. Am
J Physiol Heart Circ Physiol 2008;294:H456–H465
43. St-Pierre J, Buckingham JA, Roebuck SJ, Brand MD. Topology of superoxide production from different sites in the mitochondrial electron
transport chain. J Biol Chem 2002;277:44784–44790
44. Seifert EL, Estey C, Xuan JY, Harper ME. Electron transport chaindependent and -independent mechanisms of mitochondrial H2O2 emission
during long-chain fatty acid oxidation. J Biol Chem 2010;285:5748–5758
45. Du X, Edelstein D, Obici S, Higham N, Zou MH, Brownlee M. Insulin resistance reduces arterial prostacyclin synthase and eNOS activities by
increasing endothelial fatty acid oxidation. J Clin Invest 2006;116:1071–
1080
46. Yamagishi SI, Edelstein D, Du XL, Kaneda Y, Guzmán M, Brownlee M.
Leptin induces mitochondrial superoxide production and monocyte chemoattractant protein-1 expression in aortic endothelial cells by increasing
fatty acid oxidation via protein kinase A. J Biol Chem 2001;276:25096–
25100
47. Muller FL, Liu Y, Van Remmen H. Complex III releases superoxide to both
sides of the inner mitochondrial membrane. J Biol Chem 2004;279:49064–
49073
48. Hall CL, Lambeth JD. Studies on electron transfer from general acyl-CoA
dehydrogenase to electron transfer flavoprotein. J Biol Chem 1980;255:
3591–3595
49. Steenkamp DJ, Husain M. The effect of tetrahydrofolate on the reduction
of electron transfer flavoprotein by sarcosine and dimethylglycine dehydrogenases. Biochem J 1982;203:707–715
50. Zhang J, Frerman FE, Kim JJ. Structure of electron transfer flavoproteinubiquinone oxidoreductase and electron transfer to the mitochondrial
ubiquinone pool. Proc Natl Acad Sci USA 2006;103:16212–16217
diabetes.diabetesjournals.org